Picosecond-Resolved Fluorescent Probes at Functionally Distinct

Jun 3, 2014 - Corey W. Meadows , Gurusamy Balakrishnan , Brandon L. Kier , Thomas G. ... Corey W. Meadows , Jonathan E. Tsang , and Judith P. Klinman...
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Picosecond-Resolved Fluorescent Probes at Functionally Distinct Tryptophans within a Thermophilic Alcohol Dehydrogenase: Relationship of Temperature-Dependent Changes in Fluorescence to Catalysis Corey W. Meadows,†,§ Ryan Ou,† and Judith P. Klinman*,†,‡,§ †

Department of Chemistry, ‡Department of Molecular and Cell Biology, and the §California Institute for Quantitative Biosciences, University of California, Berkeley, Berkeley, California 94720, United States S Supporting Information *

ABSTRACT: Two single-tryptophan variants were generated in a thermophilic alcohol dehydrogenase with the goal of correlating temperature-dependent changes in local fluorescence with the previously demonstrated catalytic break at ca. 30 °C (Kohen et al., Nature 1999, 399, 496). One tryptophan variant, W87in, resides at the active site within van der Waals contact of bound alcohol substrate; the other variant, W167in, is a remote-site surface reporter located >25 Å from the active site. Picosecond-resolved fluorescence measurements were used to analyze fluorescence lifetimes, time-dependent Stokes shifts, and the extent of collisional quenching at Trp87 and Trp167 as a function of temperature. A subnanosecond fluorescence decay rate constant has been detected for W87in that is ascribed to the proximity of the active site Zn2+ and shows a break in behavior at 30 °C. For the remainder of the reported lifetime measurements, there is no detectable break between 10 and 50 °C, in contrast with previously reported hydrogen/deuterium exchange experiments that revealed a temperature-dependent break analogous to catalysis (Liang et al., Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 9556). We conclude that the motions that lead to the rigidification of ht-ADH below 30 °C are likely to be dominated by global processes slower than the picosecond to nanosecond motions measured herein. In the case of collisional quenching of fluorescence by acrylamide, W87in and W167in behave in a similar manner that resembles free tryptophan in water. Stokes shift measurements, by contrast, show distinctive behaviors in which the active-site tryptophan relaxation is highly temperature-dependent, whereas the solvent-exposed tryptophan’s dynamics are temperature-independent. These data are concluded to reflect a significantly constrained environment surrounding the active site Trp87 that both increases the magnitude of the Stokes shift and its temperature-dependence. The results are discussed in the context of spatially distinct differences in enthalpic barriers for protein conformational sampling that may be related to catalysis.

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INTRODUCTION Understanding the role that protein dynamics play in enzyme catalysis is an area of intense research efforts and vigorous debate.1−6 Because timescales for protein motions range from femtoseconds (fs)/picoseconds (ps) to seconds,7 multiple kinetic and spectroscopic approaches are needed to interrogate the roles of slow versus fast motions in reaction rate acceleration. Historically, the analysis of kinetic isotope effects (KIEs) has contributed significantly to our understanding of the contribution of proximal and distal motions during C−H bond activation.8−13 Such analyses have included secondary KIEs,14,15 while being more generally associated with the properties of primary hydrogen KIEs and their temperature dependencies.13 Secondary KIEs have also been extended to implicate protein dynamics during heavy atom group transfer reactions.16,17 As spectroscopic techniques have been developed for macromolecular systems such as enzymes, there has been an explosion in the diversity of techniques used to probe © XXXX American Chemical Society

protein dynamics. Namely, advances in techniques involving nuclear magnetic resonance (NMR),7,18−20 hydrogen/deuterium (H/D) exchange mass spectrometry,21−23 X-ray crystallography,24,25 single molecule methods,26−28 temperature-jump methods,29,30 and vibrational spectroscopy,31−33 are but a few of the experimental approaches available to characterize virtually any timescale of protein motion. However, obtaining evidence that relates a specific protein motion to the variables that control function is a much more elusive goal. In the latter context, many ideas have emerged, with the goal of linking a protein’s catalytic efficiency to the extensive degrees of freedom accessible within these large systems. Proposals put forth include the involvement of networks of protein motions,2 the creation of near-attack conformers,34 rate-promoting Received: January 23, 2014 Revised: May 5, 2014

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protein modes,35 and protein conformational selection.7,36 The temperature-dependence of local probes of protein dynamics and electrostatics can be particularly informative and has been implemented via measurements of the Stark shift,37,38 Stokes shift,39−41 and T-jump fluorescence methods.42 Most of the completed studies have been implemented in the context of very low temperatures, characterizing features of the glass transition within proteins,43 or the impact of varying solvents.44 Herein, we provide a rigorous study that analyzes dynamical behavior at distinct functional locales within ht-ADH in the context of two functionally disparate temperature regimes. Time-resolved fluorescence techniques, focusing either on the intrinsic fluorescence of tryptophan side chains or site specifically incorporated unnatural amino acids, are increasingly employed to measure fluorescence lifetimes, the time dependence of Stokes shifts, and the time constants for fluorescence quenching in the presence of exogenous ligands. Of particular interest is the time dependence of the reorganization of the surrounding medium in response to excitation at a single fluorescent side chain through the construction of timeresolved emission spectra (TRES), and the quantification of the corresponding time-dependent red shifts. Early investigations of time-dependent Stokes shifts measured the responses of homogeneous solvents of varying polarity to excited-state dipoles created in small molecules.45−48 In light of the recent increased interest in protein dynamics, timedependent Stokes shifts have also been pursued in proteins to better understand structure−function relationships,41,49−51 rotameric interconversions,52,53 the role of biological water layers,54−57 and enzyme reaction coordinates.58 Moreover, Stern−Volmer analyses have been beneficial in understanding degrees of amino acid solvent exposure in proteins,51,59,60 interactions of quenchers with fluorophores,61,62 and characterization of protein conformational changes.63,64 Decades of kinetic and mass spectrometric data collected on a thermophilic alcohol dehydrogenase (ht-ADH) isolated from Bacillus stearothermophilus (B. stearothermophilus) have led to an integrated model that relates the properties of hydride transfer to the inherent properties of the conformational ensemble accessible to the enzyme. Arrhenius plots of the temperaturedependent hydride transfer within ht-ADH exhibit a discontinuous “break” at 30 °C in which the enthalpy of activation decreases by ca. 7 kcal/mol at higher temperatures (i.e., 22 kcal/mol < 30 °C; 15 kcal/mol > 30 °C).8 Moreover, the KIE data reveal that hydride transfer is temperaturedependent below 30 °C and temperature-independent above 30 °C. The aggregate kinetic data imply that the same enzyme can achieve two drastically different conformational ensembles with distinctive tunneling properties. Structural insight into the two temperature regimes of ht-ADH has come from the time and temperature-dependence of H/D exchange within the apoenzyme.22 Among twenty-one peptides analyzed in htADH, representing almost full coverage of the protein sequence, five peptides were found to exhibit increased amide backbone solvent exchange with D2O with a transition temperature of 40−55 °C. Another five peptides located exclusively within the substrate-binding domain showed increased protein backbone deuteron incorporation, transitioning between 20−40 °C. These H/D exchange experiments in apoprotein have yielded a spatial resolution of the temperaturedependent change in protein flexibility that correlates with the catalytic behavior of ht-ADH. More recently, the impact of mutation of two cofactor binding domain residues (Leu176 and

Val260) to side chains of reduced mass has led to a model in which the relative population of rigid, catalytically incompetent microstates and more flexible, tunneling-ready microstates govern not only the tunneling properties but also the magnitude of the experimental enthalpies and entropies of activation.65 The present work utilizes ps-resolved fluorescence spectroscopy to investigate fluorescence lifetimes, the temperaturedependence of time-dependent Stokes shifts, and collisional quenching of ht-ADH by acrylamide. In order to observe the enzyme dynamics with minimal perturbation, the red-edge excitation effect of tryptophan was exploited to minimize contributions from fluorescent tyrosines and phenylalanines.66 The native wild-type (WT) ht-ADH monomeric structure contains three tryptophans. Because lifetime measurements of WT protein cannot distinguish the origin of fluorescence without a priori knowledge about each locale, single-tryptophan variants were created such that two of the three tryptophans were replaced with combinations of either tyrosine or phenylalanine. Of the three single-Trp possibilities, two of them were found to have kinetic properties that closely mirror WT behavior. These constructs are W49F:W167Y (W87in), which fortuitously resides at the enzyme active site within van der Waals contact of the substrate-binding pocket, and W49F:W87F (W167in), a solvent-exposed, remote-site tryptophan found over 25 Å away from the catalytic center (Figure 1).

Figure 1. Structure of monomeric ht-ADH (PDB: 1RJW). The relative locations of the two tryptophans (red) and substrate analogue trifluoroethanol (black) are represented with sticks. The catalytic zinc inside the active site (gray) is represented as a sphere.

Expanded representations of the side chains surrounding each tryptophan are shown in Figure 2. As will be presented herein, fluorescence lifetimes and time-dependent Stokes shifts reveal a greater sensitivity to environmental differences at each single tryptophan site than their collisional quenching behaviors. The data are discussed in the context of the unique properties of the W87in as they relate to catalysis.

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EXPERIMENTAL SECTION Site-Directed Mutagenesis and Protein Purification. W87in (W49F:W167Y) and W167in (W49F:W87F) were generated by following protocols in the QuikChange sitedirected mutagenesis kit (Agilent Technologies). Mutant plasmids were generated and amplified using a PTC-2000 Peltier Thermal Cycler (MJ Research). Appropriate combina-

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Figure 2. Closeups of residues surrounding Trp87 (left) and Trp167 (right). Tryptophans are shown in red. Residues with side-chain interactions within 6 Å of the tryptophan are shown for each locale. Amide backbone atoms were omitted for clarity.

appropriately diluted enzyme. Initial rates were determined by measuring the slopes of the linear traces acquired for at least 3 min. Such conditions for turnover were assayed over nine temperatures ranging from 7−51.5 °C. Assays were incubated in a water bath before placement into the spectrophotometer’s Peltier-controlled cuvette cell. Kinetic parameters were obtained from a nonlinear fit to the 2D-Michaelis−Menten equation using DataFit (Oakdale Engineering); the corresponding activation parameters were obtained from nonlinear fits to the Arrhenius equation using DataFit. Time-Dependent Activity Loss. The activity loss of W87in and W167in were tested under conditions that would reflect the stability of the protein spanning the duration of timeresolved fluorescence measurements (typically 2−3 h for measurements of 7 wavelengths at each temperature). Each respective variant was diluted to a concentration of 3−4 μM and incubated for 4 h at 10 °C, 30 °C, and 50 °C. The activity was measured under saturating conditions (10 mM NAD+, 15 mM d-BnOH) at ten time points ranging from 2 min to 4 h after enzyme incubation in triplicate. Excessive activity loss is defined as leading to less than 75% of initial activity after a 2 min incubation. For W87in, there was little difference in protein stability profiles at 10 and 30 °C, while enzyme inactivation is somewhat faster at 50 °C (Figure S2 in the Supporting Information). On the basis of the observations, W87in samples were changed after 150 min of measurement between 10 and 30 °C and every 90 min at temperatures above 30 °C. For W167in, the protein stability profiles indicated a smaller temperature-dependence at low temperatures. W167in does not show excessive loss of activity at 10 or 30 °C over a 4 h incubation period, although it is less stable than W87 in a high temperature (Figure S2 in the Supporting Information). For W167in, samples were not changed for lifetime measurements collected between 10−30 °C and replaced every 75 min at temperatures above 30 °C. Other Controls. The oligomeric state and secondary structure of the single-Trp mutants were verified by sizeexclusion chromatography and CD, respectively, using methods previously described.66 Circular dichroism (CD) spectra comparing W87in and W167in to the WT secondary structure are shown at 30 °C (Figure S3 in the Supporting Information). No major changes in secondary structure were observed across all variants relative to WT, verifying that the conservative substitutions contained within each variant do not cause a large change in overall protein structure. Gel filtration chromatograms are also shown at 4 °C (Figure S4 in the Supporting Information). Both variants elute predominantly as a tetramer,

tions of the following primers (Operon) and their respective reverse complements (not shown) were introduced to the PCR mixtures to generate the single tryptophan variants: W49F: 5′CCGCTCACGGCGATTTCCCGGTAAAACCAAAAC-3′; W87F: 5′-CCGCGTTGGAATTCCTTTCTTATATTCTGCATGC-3′; W167Y: 5′-GTAACAGGGGCAAAACCAGGAGAATATGTAGCAATTTACGGTAT-3′. Changes to the pET-24b(+) Escherichia coli expression vector containing the B. stearothermophilus ht-ADH gene are underlined. Gene sequencing was performed by the UC Berkeley DNA Sequencing Facility. Plasmid purification for gene sequencing was executed using the materials and protocols contained within the QIAprep spin miniprep kit (QIAGEN). Tyrosine was chosen for position 167 due to significant activity loss (ca. 70%) after 30 min of incubation at 4 °C for W49F:W167F. Subsequent plasmid transformation, cell growth, and lysis are described elsewhere.22 The single-tryptophan variants were purified as previously described with some minor modifications.67 Because bound NADH quenches Trp87 fluorescence via FRET (Figure S1 in the Supporting Information), 70 mL of 1 mM adenosine 5′monophosphate (Sigma-Aldrich) was used to elute the single tryptophan variants from the 5′-AMP Agarose affinity column. This was used in lieu of NAD+ to avoid any potential distortion of fluorescence measurements. After affinity column elution, the protein was spin concentrated to 8 Å from the closest approach to bulk solvent, rendering it too far within the protein core to have random long-range interactions that distort the quenching behavior. Regardless of whether quenching is occurring as the result of direct collision or via some through-space interaction, we expected that fluctuations of the protein were likely to be involved in transport of the acrylamide to a distance close enough to facilitate fluorescence quenching. Nonetheless, the effect of temperature on the Stern−Volmer data reveals a very similar temperature-dependence at both positions. The two variants display a ΔH‡ close to 4 kcal/mol, consistent with the activation energy measured for acrylamide quenching of free Nacetyl-tryptophanamide in water.72 This further corroborates the notion that Trp87 near the catalytic center has a similar degree of exposure to solvent as Trp167. The most compelling observation from the quenching data at both sites is the lack of an enthalpic transition at 30 °C. Both Trp87 and Trp167 belong to peptides that exhibit increased solvent exchange above 30 °C as detected by H/D exchange under the EX2 condition.22 While this H/D exchange occurs over slow timescales, a hierarchy of motions may be expected to contribute to the local unfolding that allows transient access of the solvent D2O/OD− to the protein peptide backbone. The fact that collisional quenching of fluorescence is insensitive to

Figure 7. Arrhenius plots of the Stokes shift decay constants for W87in (red) and W167in (blue).

Figure 8. Collisional Stern−Volmer plots for W87in (top) and W167in (bottom). The representative temperatures are shown at 10 °C (black), 20 °C (blue), 30 °C (green), 40 °C (orange), and 50 °C (red).

Figure 9. Arrhenius plots of the bimolecular quenching constants derived from the slopes plotted in Figure 9 for W87in (red) and W167in (blue).

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experiments. The presence of a very modest Stokes shift in the nanosecond regime for W167in may be due to a large number of van der Waals interactions (r‑6 dependence) in the proximity of this side chain. By contrast, W87in has a broad diversity of side chains interacting near the indole moiety. The catalytic Zn2+ with coordinated Cys148 and His61 are excellent candidates to quench inherent fluorescent lifetimes and are likely the origin of the sub-nanosecond component in the triexponential fluoresence lifetime analysis (Table 2). Asn111 and Gln109 are excellent hydrogen-bonding candidates that contain heavy atoms measured within 4 Å of the indole nitrogen. Gln109, Tyr114, and Ala112 also contribute amide backbone interactions to Trp87. This extremely heterogeneous array of interactions surrounding Trp87 can easily explain the large enthalpy of greater than 9 kcal/mol for the excited-state reorganization. The catalytic center is 8 Å from the bulk solvent, resulting in lower dielectric constants that may be expected to require a high-energy input to break the combination of ionic and dipole−dipole interactions (r‑2 and r‑3 dependence). Relationship of the Fluorescence Data for ht-ADH to Kinetic and H/D Exchange Behavior. The spatial resolution of the Stokes shifts investigated for ht-ADH herein allows comparison to an earlier study of the temperature-dependence of H/D exchange within a thermophilic dihydrofolate reductase (DHFR).21 Despite the very different timescales of fluorescence and deuterium exchange studies, DHFR showed a similar pattern of a large local enthalpy (for unfolding) near the cofactor and substrate-binding site that contrasted with very small enthalpies (for local unfolding) at regions distant from the active site and near the protein surface. The studies of native DHFR led to the proposal of a largely entropy-driven local unfolding throughout the majority of the protein, which is the result of small enthalpic barriers for the interconversion of multiple conformers and reflects instead the probability of reaching a state amenable to solvent penetration and backbone exchange. The active site on the other hand, with its large number of specific interactions, was proposed to require substantial thermal activation to unfold sufficiently to expose backbone amides to the solvent.21 The common theme between the experiments with DHFR and the present study is the existence of extensive structure at the active site that is dominated by highly specific and concentrated dipolar and electrostatic interactions. These interactions are also likely to be the primary cause of the reorganization energy that is required to tune the electrostatics of the hydrogen donor and acceptor in C−H activation reactions.13 While the value of the enthalpy of activation for the Stokes shift at W87in for ht-ADH is approximately half of the barrier measured on catalysis above 30 °C, it is reasonable to assume that there is overlap among the environmental changes that affect both processes. Among all of the fluorescent measurements on ht-ADH, the single one that shows any possible break with temperature analogous to catalysis and H/D exchange is a small amplitude sub-nanosecond fluorescence decay at Trp87 (Table 2 and Figure 4). This is completely absent from Trp167 and is ascribed to an interaction between Trp87 and the active site Zn2+. The arrangement of ligands to the active site metal catalyst may be expected to be very sensitive to temperaturedependent structural changes that likewise alter catalysis. The remainder of the fluorescent methods reflect motions than can be attributed to side chain rotations, backbone fluctuations,

the temperature transition that controls H/D exchange (and catalysis) could mean either that the timescale of motions controlling the fluorescence properties is not relevant or that the large degree of solvent exposure at both Trp87 and Trp167 results in an insensitivity of the quenching parameters to nanosecond motions. The Stokes shift data become especially relevant and informative in the latter context. Temperature-Dependent Stokes Shifts Reveal Opposing Temperature-Dependent Effects at W87in and W167in. In general, the time-dependent Stokes shift reports on the evolution of the solvent’s electrostatic environment during the excited state of a fluorophore. Instantly after excitation, tryptophan is in its most electronically unfavorable environment because the surrounding residues have not yet rearranged to accommodate the newly generated electronic distortion, in accordance with the Franck−Condon (FC) principle. Hence, the TRES at τfc−1 is calculated to be the highest energy state for emission and is referred to as the FC state. During the excited state, the solvent reorganizes at a rate constant, ksol, producing more energetically favorable TRES with peaks shifting to red wavelengths until the steady-state emission wavelength at time τr−1 is reached.69−71 The temperature-dependence of the red shift decay rates reveals two opposing behaviors, featuring an active site environment at Trp87 that is highly temperature-dependent and a surface environment at Trp167 that is practically temperature-independent (Figure 7). This is accompanied by very different emission maxima at t = 0, with the W167in showing a much more red-shifted spectrum at the initial time point (Table 3). These two features are proposed to be intimately related to one another and suggest that significant amount of rearrangement at Trp167 is occurring on a much faster timescale than nanoseconds at all temperatures (cf. Table 4). Studies employing fluorescence upconversion techniques with femtosecond time resolution can report on distortions caused by water reorganization and water-coupled protein interactions having relaxation timescales on the order of 102− 104 fs. Such solvation decays indicate much more drastic Stokes shifts of tryptophan fluorescence, relaxing nearly 800−1800 cm−1 to the red of their respective FC states.54−57 The fact that the overall shift for W87in is 486 cm−1 at 10 °C, in comparison to 100 cm−1 for W167in at the same temperature, is striking and shows that the environment at W87in is much more restricted, requiring an input of thermal energy before it can approach the behavior of W167in at 50 °C. The origin of the factors controlling the respective enthalpic barriers can be rationalized according to the local structure around each tryptophan (Figure 2). Trp167 is slightly more accessible to solvent and resides in a homogeneous environment relative to Trp87. In fact, only two polar interactions exist near Trp167 (Figure 2). The side chain of Asn190 is a hydrogen-bonding partner with the indole nitrogen. His232 is the only other polar side chain interaction within 6 Å of Trp167 but is poorly aligned for any polar interactions because its imidazole moiety lies coplanar with the indole moiety. The sole Asn190 interaction with Trp167 is most likely responsible for stabilizing the excited state tryptophan, as the ca. 6 nanosecond lifetime is more than twice the value (2.7 ns) measured for free Trp in water.80 However, because of the scarce number of polar interactions at this site, the environment is unfavorable for accommodation of an excited-state fluorophore relative to the active site. This feature is proposed to result in a predominant relaxation at a faster rate than the time resolution of the present J

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loop fluctuations, and translations of side chains.7,51,81 Motions of this type and timescale are functionally quite local and in the vicinity of Trp87 do not appear to correlate with the previously demonstrated transition in tunneling properties at 30 °C. We conclude that the nanosecond to picosecond motions detected from fluorescence measurements are likely spatially distinct from the similar time scale motions that control donor−acceptor distance sampling of the hydrogen tunneling coordinate under various conditions. Further, the latter motions may only be detectable in binary and tertiary complexes of enzymes. By contrast, the specific protein motions that generate a reversible trapping of ht-ADH into low enthalpy microstates below 30 °C are evident in τ1 for W87in (Table 2 and Figure 4), analogous to transitions seen earlier via H/D exchange. These properties point toward longer and more global motions controlling the transitions seen at 30 °C, warranting future studies of dynamics in ht-ADH on the microsecond timescale. Such motions may be directly linked to protein subunit interactions, as implicated from recent studies that show a connection between a Tyr25 side chain at the dimer interface of ht-ADH that dictates the presence of the Arrhenius break and the region of the active site where Trp87 resides.82 Extension of spectroscopic studies to mutant forms of ht-ADH known to profoundly alter the properties of the catalytically linked temperature transition are also likely to be quite informative.

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Unfolding Mechanism for Allostery and Functional Adaptation in Proteins. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 16984−16989. (5) Kamerlin, S. C. L.; Warshel, A. At the Dawn of the 21st Century: Is Dynamics the Missing Link for Understanding Enzyme Catalysis? Proteins: Struct., Funct., Bioinf. 2010, 78, 1339−1375. (6) Forconi, M.; Porecha, R. H.; Piccirilli, J. A.; Herschlag, D. Tightening of Active Site Interactions En Route to the Transition State Revealed by Single-Atom Substitution in the Guanosine-Binding Site of the Tetrahymena Group I Ribozyme. J. Am. Chem. Soc. 2011, 133, 7791−7800. (7) Henzler-Wildman, K. A.; Lei, M.; Thai, V.; Kerns, S. J.; Karplus, M.; Kern, D. A Hierarchy of Timescales in Protein Dynamics is Linked to Enzyme Catalysis. Nature 2007, 450, 913−916. (8) Kohen, A.; Cannio, R.; Bartolucci, S.; Klinman, J. P. Enzyme Dynamics and Hydrogen Tunneling in a Thermophilic Alcohol Dehydrogenase. Nature 1999, 399, 496−499. (9) Knapp, M. J.; Rickert, K.; Klinman, J. P. Temperature-Dependent Isotope Effects in Soybean Lipoxygenase-1: Correlating Hydrogen Tunneling with Protein Dynamics. J. Am. Chem. Soc. 2002, 124, 3865− 3874. (10) Hatcher, E.; Soudackov, A. V.; Hammes-Schiffer, S. ProtonCoupled Electron Transfer in Soybean Lipoxygenase: Dynamical Behavior and Temperature Dependence of Kinetic Isotope Effects. J. Am. Chem. Soc. 2007, 129, 187−196. (11) Maglia, G.; Allemann, R. K. Evidence for Environmentally Coupled Hydrogen Tunneling During Dihydrofolate Reductase Catalysis. J. Am. Chem. Soc. 2003, 125, 13372−13373. (12) Pudney, C. R.; Johannissen, L. O.; Sutcliffe, M. J.; Hay, S.; Scrutton, N. S. Direct Analysis of Donor-Acceptor Distance and Relationship to Isotope Effects and the Force Constant for Barrier Compression in Enzymatic H-Tunneling Reactions. J. Am. Chem. Soc. 2010, 132, 11329−11335. (13) Klinman, J. P.; Kohen, A. Hydrogen Tunneling Links Protein Dynamics to Enzyme Catalysis. Annu. Rev. Biochem. 2013, 82, 471− 496. (14) Cha, Y.; Murray, C. J.; Klinman, J. P. Hydrogen Tunneling in Enzyme Reactions. Science 1989, 243, 1325−1330. (15) Roston, D.; Kohen, A. Elusive Transition State of Alcohol Dehydrogenase Unveiled. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 9572−9577. (16) Zhang, J. Y.; Klinman, J. P. Enzymatic Methyl Transfer: Role of an Active Site Residue in Generating Active Site Compaction That Correlates with Catalytic Efficiency. J. Am. Chem. Soc. 2011, 133, 17134−17137. (17) Schwartz, P. A.; Vetticatt, M. J.; Schramm, V. L. Transition State Analysis of the Arsenolytic Depyrimidination of Thymidine by Human Thymidine Phosphorylase. Biochemistry 2011, 50, 1412−1420. (18) Schnell, J. R.; Dyson, H. J.; Wright, P. E. Effect of Cofactor Binding and Loop Conformation on Side Chain Methyl Dynamics in Dihydrofolate Reductase. Biochemistry 2004, 43, 374−383. (19) Baldwin, A. J.; Kay, L. E. NMR Spectroscopy Brings Invisible Protein States into Focus. Nat. Chem. Biol. 2009, 5, 808−814. (20) Osborne, M. J.; Schnell, J.; Benkovic, S. J.; Dyson, H. J.; Wright, P. E. Backbone Dynamics in Dihydrofolate Reductase Complexes: Role of Loop Flexibility in the Catalytic Mechanism. Biochemistry 2001, 40, 9846−9859. (21) Oyeyemi, O. A.; Sours, K. M.; Lee, T.; Resing, K. A.; Ahn, N. G.; Klinman, J. P. Temperature Dependence of Protein Motions in a Thermophilic Dihydrofolate Reductase and its Relationship to Catalytic Efficiency. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 10074− 10079. (22) Liang, Z.-X.; Lee, T.; Resing, K. A.; Ahn, N. G.; Klinman, J. P. Thermal-Activated Protein Mobility and its Correlation with Catalysis in Thermophilic Alcohol Dehydrogenase. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 9556−9561. (23) Zhu, M. M.; Rempel, D. L.; Du, Z. H.; Gross, M. L. Quantification of Protein-Ligand Interactions by Mass Spectrometry, Titration, and H/D Exchange: PLIMSTEX. J. Am. Chem. Soc. 2003, 125, 5252−5253.

ASSOCIATED CONTENT

* Supporting Information S

Steady-state fluorescence spectra, Stern−Volmer rate constants, enzyme kinetics, CD spectra, and gel-filtration data. This material is available free of charge via the Internet at http:// pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Tel: (510) 642-2668. Fax: (510) 643-4500. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was funded by a grant from the National Institutes of Health to J.P.K. (GM025765 and GM025765-32S1). ABBREVIATIONS: NAD+, nicotinamide adenine dinucleotide; ht-ADH, thermophilic alcohol dehydrogenase from Bacillus stearothermophilus; KIE, kinetic isotope effect; WT, wild-type; TRES, time-resolved emission spectra; TCSPC, time-correlated single photon counting; BnOH, benzyl alcohol

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REFERENCES

(1) Nagel, Z. D.; Klinman, J. P. Update 1 of: Tunneling and Dynamics in Enzymatic Hydride Transfer. Chem. Rev. 2010, 110, Pr41−Pr67. (2) Hammes-Schiffer, S.; Benkovic, S. J. Relating Protein Motion to Catalysis. Annu. Rev. Biochem. 2006, 75, 519−41. (3) Schwartz, S.; Schramm, V. L. Enzymatic Transition States and Dynamic Motion in Barrier Crossing. Nat. Chem. Biol. 2009, 5, 551− 558. (4) Schrank, T. P.; Bolen, D. W.; Hilser, V. J. Rational Modulation of Conformational Fluctuations in Adenylate Kinase Reveals a Local K

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Article

(24) Fraser, J. S.; Clarkson, M. W.; Degnan, S. C.; Erion, R.; Kern, D.; Alber, T. Hidden Alternative Structures of Proline Isomerase Essential for Catalysis. Nature 2009, 462, 669−673. (25) Schotte, F.; Lim, M. H.; Jackson, T. A.; Smirnov, A. V.; Soman, J.; Olson, J. S.; Phillips, G. N.; Wulff, M.; Anfinrud, P. A. Watching a Protein as it Functions with 150-ps Time-Resolved X-ray Crystallography. Science 2003, 300, 1944−1947. (26) Flynn, E. M.; Hanson, J. A.; Alber, T.; Yang, H. Dynamic ActiveSite Protection by the M. tuberculosis Protein Tyrosine Phosphatase PtpB Lid Domain. J. Am. Chem. Soc. 2010, 132, 4772−4780. (27) Min, W.; English, B. P.; Luo, G. B.; Cherayil, B. J.; Kou, S. C.; Xie, X. S. Fluctuating Enzymes: Lessons from Single-Molecule Studies. Acc. Chem. Res. 2005, 38, 923−931. (28) Yang, H.; Luo, G.; Karnchanaphanurach, P.; Louie, T.-M.; Rech, I.; Cova, S.; Xun, L.; Xie, X. S. Protein Conformational Dynamics Probed by Single-Molecule Electron Transfer. Science 2003, 302, 262− 266. (29) Phillips, R. S.; Miles, E. W.; McPhie, P.; Marchal, S.; Georges, C.; Dupont, Y.; Lange, R. Pressure and Temperature Jump Relaxation Kinetics of the Conformational Change in Salmonella typhimurium Tryptophan Synthase L-Serine Complex: Large Activation Compressibility and Heat Capacity Changes Demonstrate the Contribution of Solvation. J. Am. Chem. Soc. 2008, 130, 13580−13588. (30) Ye, M. P.; Zhang, Q. L.; Li, H.; Weng, Y. X.; Wang, W. C.; Qiu, X. G. Infrared Spectroscopic Discrimination between the Loop and Alpha-Helices and Determination of the Loop Diffusion Kinetics by Temperature-Jump Time-Resolved Infrared Spectroscopy for Cytochrome c. Biophys. J. 2007, 93, 2756−2766. (31) Balakrishnan, G.; Hu, Y.; Oyerinde, O. F.; Su, J.; Groves, J. T.; Spiro, T. G.; Conformational, A. Switch to Beta-Sheet Structure in Cytochrome c Leads to Heme Exposure. Implications for Cardiolipin Peroxidation and Apoptosis. J. Am. Chem. Soc. 2007, 129, 504−505. (32) Jha, S. K.; Ji, M. B. A.; Gaffney, K. J.; Boxer, S. G. Direct Measurement of the Protein Response to an Electrostatic Perturbation that Mimics the Catalytic Cycle in Ketosteroid Isomerase. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 16612−16617. (33) Bandaria, J. N.; Dutta, S.; Nydegger, M. W.; Rock, W.; Kohen, A.; Cheatum, C. M. Characterizing the Dynamics of Functionally Relevant Complexes of Formate Dehydrogenase. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 17974−17979. (34) Hur, S.; Bruice, T. C. The Near Attack Conformation Approach to the Study of the Chorismate to Prephenate Reaction. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 12015−12020. (35) Mincer, J. S.; Schwartz, S. D. Rate-Promoting Vibrations and Coupled Hydrogen-Electron Transfer Reactions in the Condensed Phase: A Model for Enzymatic Catalysis. J. Chem. Phys. 2004, 120, 7755−7760. (36) Boehr, D. D.; Nussinov, R.; Wright, P. E. The Role of Dynamic Conformational Ensembles in Biomolecular Recognition. Nat. Chem. Biol. 2009, 5, 789−796. (37) Suydam, I. T.; Snow, C. D.; Pande, V. S.; Boxer, S. G. Electric Fields at the Active Site of an Enzyme: Direct Comparison of Experiment with Theory. Science 2006, 313, 200−204. (38) Fafarman, A. T.; Sigala, P. A.; Schwans, J. P.; Fenn, T. D.; Herschlag, D.; Boxer, S. G. Quantitative, Directional Measurement of Electric Field Heterogeneity in the Active Site of Ketosteroid Isomerase. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, E299−E308. (39) Chang, C. W.; He, T. F.; Guo, L. J.; Stevens, J. A.; Li, T. P.; Wang, L. J.; Zhong, D. P. Mapping Solvation Dynamics at the Function Site of Flavodoxin in Three Redox States. J. Am. Chem. Soc. 2010, 132, 12741−12747. (40) Stevens, J. A.; Link, J. J.; Kao, Y. T.; Zang, C.; Wang, L. J.; Zhong, D. P. Ultrafast Dynamics of Resonance Energy Transfer in Myoglobin: Probing Local Conformation Fluctuations. J. Phys. Chem. B 2010, 114, 1498−1505. (41) Abbyad, P.; Shi, X. H.; Childs, W.; McAnaney, T. B.; Cohen, B. E.; Boxer, S. G. Measurement of Solvation Responses at Multiple Sites in a Globular Protein. J. Phys. Chem. B 2007, 111, 8269−8276.

(42) Nie, B. N.; Deng, H.; Desamero, R.; Callender, R. Large Scale Dynamics of the Michaelis Complex in Bacillus stearothermophilus Lactate Dehydrogenase Revealed by a Single-Tryptophan Mutant Study. Biochemistry 2013, 52, 1886−1892. (43) Abbyad, P.; Childs, W.; Shi, X. H.; Boxer, S. G. Dynamic Stokes Shift in Green Fluorescent Protein Variants. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 20189−20194. (44) Litvinenko, K. L.; Webber, N. M.; Meech, S. R. Internal Conversion in the Chromophore of the Green Fluorescent Protein: Temperature Dependence and Isoviscosity Analysis. J. Phys. Chem. A 2003, 107, 2616−2623. (45) Chakrabarti, S. K.; Ware, W. R. Nanosecond Time-Resolved Emission Spectroscopy of 1-Anilino-8-Naphthalene Sulfonate. J. Chem. Phys. 1971, 55, 5494−5498. (46) Bagchi, B.; Oxtoby, D. W.; Fleming, G. R. Theory of the Time Development of the Stokes Shift in Polar Media. Chem. Phys. 1984, 86, 257−267. (47) Ruggiero, A. J.; Todd, D. C.; Fleming, G. R. Subpicosecond Fluorescence Anisotropy Studies of Tryptophan in Water. J. Am. Chem. Soc. 1990, 112, 1003−1014. (48) Jimenez, R.; Fleming, G. R.; Kumar, P. V.; Maroncelli, M. Femtosecond Solvation Dynamics of Water. Nature 1994, 369, 471− 473. (49) Chen, J. J.; Toptygin, D.; Brand, L.; King, J. Mechanism of the Efficient Tryptophan Fluorescence Quenching in Human Gamma DCrystallin Studied by Time-Resolved Fluorescence. Biochemistry 2008, 47, 10705−10721. (50) Guha, S.; Sahu, K.; Roy, D.; Mondal, S. K.; Roy, S.; Bhattacharyya, K. Slow Solvation Dynamics at the Active Site of an Enzyme: Implications for Catalysis. Biochemistry 2005, 44, 8940−8947. (51) Jha, A.; Ishii, K.; Udgaonkar, J. B.; Tahara, T.; Krishnamoorthy, G. Exploration of the Correlation between Solvation Dynamics and Internal Dynamics of a Protein. Biochemistry 2011, 50, 397−408. (52) Ferreira, S. T. Fluorescence Studies of the Conformational Dynamics of Parvalbumin in Solution Lifetime and Rotational Motions of the Single Tryptophan Residue. Biochemistry 1989, 28, 10066− 10072. (53) Maglia, G.; Jonckheer, A.; De Maeyer, M.; Frere, J. M.; Engelborghs, Y. An Unusual Red-Edge Excitation and Time-Dependent Stokes Shift in the Single Tryptophan Mutant Protein DDCarboxypeptidase from Streptomyces: The Role of Dynamics and Tryptophan Rotamers. Protein Sci. 2008, 17, 352−361. (54) Qiu, W. H.; Kao, Y. T.; Zhang, L. Y.; Yang, Y.; Wang, L. J.; Stites, W. E.; Zhong, D. P.; Zewail, A. H. Protein Surface Hydration Mapped by Site-Specific Mutations. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 13979−13984. (55) Qiu, W. H.; Wang, L. J.; Lu, W. Y.; Boechler, A.; Sanders, D. A. R.; Zhong, D. P. Dissection of Complex Protein Dynamics in Human Thioredoxin. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 5366−5371. (56) Zhang, L. Y.; Wang, L. J.; Kao, Y. T.; Qiu, W. H.; Yang, Y.; Okobiah, O.; Zhong, D. P. Mapping Hydration Dynamics Around a Protein Surface. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 18461−18466. (57) Kwon, O. H.; Yoo, T. H.; Othona, C. M.; Van Deventer, J. A.; Tirrell, D. A.; Zewail, A. H. Hydration Dynamics at Fluorinated Protein Surfaces. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 17101−17106. (58) Childs, W.; Boxer, S. G. Solvation Response along the Reaction Coordinate in the Active Site of Ketosteroid Isomerase. J. Am. Chem. Soc. 2010, 132, 6474−6480. (59) Eftink, M. R.; Jameson, D. M. Acrylamide and Oxygen Fluorescence Quenching Studies with Liver Alcohol-Dehydrogenase Using Steady-State and Phase Fluorometry. Biochemistry 1982, 21, 4443−4449. (60) Kim, S. J.; Chowdhury, F. N.; Stryjewski, W.; Younathan, E. S.; Russo, P. S.; Barkley, M. D. Time-Resolved Fluorescence of the Single Tryptophan of Bacillus-Stearothermophilus Phosphofructokinase. Biophys. J. 1993, 65, 215−226. (61) Lakowicz, J. R.; Zelent, B.; Gryczynski, I.; Kusba, J.; Johnson, M. L. Distance-Dependent Fluorescence Quenching of Tryptophan by Acrylamide. Photochem. Photobiol. 1994, 60, 205−214. L

dx.doi.org/10.1021/jp500825x | J. Phys. Chem. B XXXX, XXX, XXX−XXX

The Journal of Physical Chemistry B

Article

(62) Strambini, G. B.; Gonnelli, M. Fluorescence Quenching of Buried Trp Residues by Acrylamide Does Not Require Penetration of the Protein Fold. J. Phys. Chem. B 2010, 114, 1089−1093. (63) Weber, J.; Senior, A. E. Features of F-1-ATPase Catalytic and Noncatalytic Sites Revealed by Fluorescence Lifetimes and Acrylamide Quenching of Specifically Inserted Tryptophan Residues. Biochemistry 2000, 39, 5287−5294. (64) Secundo, F.; Russo, C.; Giordano, A.; Carrea, G.; Rossi, M.; Raia, C. A. Temperature-induced conformational change at the catalytic site of Sulfolobus solfataricus alcohol dehydrogenase highlighted by Asn249Tyr substitution. A hydrogen/deuterium exchange, kinetic, and fluorescence quenching study. Biochemistry 2005, 44, 11040−11048. (65) Nagel, Z. D.; Dong, M.; Bahnson, B. J.; Klinman, J. P. Impaired Protein Conformational Landscapes as Revealed in Anomalous Arrhenius Prefactors. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 10520− 10525. (66) Lakowicz, J. R. Fluorescence Spectroscopy of Biomolecules. In Encyclopedia of Molecular Biology and Molecular Medicine, Meyers, R. A., Ed. VCH Publishers: New York, NY, 1995. (67) Nagel, Z. D.; Meadows, C. W.; Dong, M.; Bahnson, B. J.; Klinman, J. P. Active Site Hydrophobic Residues Impact Hydrogen Tunneling Differently in a Thermophilic Alcohol Dehydrogenase at Optimal Versus Nonoptimal Temperatures. Biochemistry 2012, 51, 4147−4156. (68) Bradford, M. M.; Rapid, A. and Sensitive Method for the Quantitation of Microgram Quantities of Protein Utilizing the Principle of Protein-Dye Binding. Anal. Biochem. 1976, 72, 248−54. (69) Badea, M. G.; Brand, L. Time-Resolved Fluorescence Measurements. Methods Enzymol. 1979, 61, 378−425. (70) Lakowicz, J. R.; Cherek, H.; Laczko, G.; Gratton, E. TimeResolved Fluorescence Emission-Spectra of Labeled PhospholipidVesicles, as Observed Using Multi-Frequency Phase-Modulation Fluorometry. Biochim. Biophys. Acta 1984, 777, 183−193. (71) Horng, M. L.; Gardecki, J. A.; Papazyan, A.; Maroncelli, M. Subpicosecond Measurements of Polar Solvation Dynamics: Coumarin-153 Revisited. J. Phys. Chem. 1995, 99, 17311−17337. (72) Eftink, M. R.; Ghiron, C. A. Exposure of Tryptophanyl Residues and Protein Dynamics. Biochemistry 1977, 16, 5546−5551. (73) Komiyama, T.; Miwa, M. Fluorescence Quenching as an Indicator for the Exposure of Tryptophyl Residues in Streptomyces Subtilisin Inhibitor. J. Biochem. 1980, 87, 1029−1036. (74) Kenoth, R.; Swamy, M. J. Steady-Sate and Time-Resolved Fluorescence Studies on Trichosanthes Cucumerina Seed Lectin. J. Photochem. Photobiol. B 2003, 69, 193−201. (75) Varnes, A. W.; Wehry, E. L.; Dodson, R. B. Interactions of Transition-Metal Ions with Photoexcited States of Flavins: Fluorescence Quenching Studies. J. Am. Chem. Soc. 1972, 94, 946−950. (76) Cook, R. L.; Langford, C. H. Metal-Ion Quenching of FulvicAcid Fluorescence Intensities and Lifetimes: Nonlinearities and a Possible 3-Component Model. Anal. Chem. 1995, 67, 174−180. (77) Tabak, M.; Sartor, G.; Neyroz, P.; Spisni, A.; Cavatorta, P. Interaction of Copper and Nickel Ions with Tryptophan and Glycyltryptophan: Time Resolved Fluorescence Study. J. Lumin. 1990, 46, 291−299. (78) Kim, S. J.; Lewis, M. S.; Knutson, J. R.; Porter, D. K.; Kumar, A.; Wilson, S. H. Characterization of the Tryptophan Fluorescence and Hydrodynamic Properties of Rat DNA-Polymerase-Beta. J. Mol. Biol. 1994, 244, 224−235. (79) Eftink, M. R.; Hagaman, K. A. Fluorescence Lifetime and Anisotropy Studies with Liver Alcohol-Dehydrogenase and Its Complexes. Biochemistry 1986, 25, 6631−6637. (80) Boens, N.; Qin, W. W.; Basaric, N.; Hofkens, J.; Ameloot, M.; Pouget, J.; Lefevre, J. P.; Valeur, B.; Gratton, E.; Vandeven, M.; et al. Fluorescence Lifetime Standards for Time and Frequency Domain Fluorescence Spectroscopy. Anal. Chem. 2007, 79, 2137−2149. (81) Boehr, D. D.; Dyson, H. J.; Wright, P. E.; An, N. M. R. Perspective on Enzyme Dynamics. Chem. Rev. 2006, 106, 3055−3079.

(82) Nagel, Z. D.; Cun, S. J.; Klinman, J. P. Identification of a Longrange Protein Network That Modulates Active Site Dynamics in Extremophilic Alcohol Dehydrogenases. J. Biol. Chem. 2013, 288, 14087−14097.

M

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